The Very Handy Manual: How to Catch and Identify Bees

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The Very Handy Manual: How to Catch and Identify Bees

and Manage a Colle
ct
ion


A Collective and Ongoing Effort by Those Who Love to Study Bees in North America

Last Revised:
May
, 2012

This manual is a compilation of the wisdom and experience of many
individuals, some of whom are directly
acknowledged here and others not. We thank all of you. The bulk of the text was compiled by Sam Droege at
the USGS Native Bee Inventory and Monitoring Lab over several years from 2004
-
2008. We regularly update
the
manual with new information, so, if you have a new technique, some additional ideas for sections,
corrections or additions, we would like to hear from you. Please email those to Sam Droege
(
sdroege@usgs.gov
). You c
an also email Sam if you are interested in joining the group’s discussion group on
bee monitoring and identification
. Many thanks to Dave and Janice Green, Tracy Zarrillo, and Liz Sellers for
their many hours of editing this manual.

"They've got this stea
mroller going, and they won't stop until
there's nobody fishing. What are they going to do then, save
some bees?"
-

Mike Russo (Massachusetts fisherman who has
fished cod for 18 years, on environmentalists)
-
Provided by
Matthew Shepherd


Contents

Where to Find Bees

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................

2

Killing Bees to Study Them

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.....

2

Ne
ts
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.........

2

Netting Technique

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..................

3

Removing Bees From the Net

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................................
.

4

Us
ing Ice and Dry Ice

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................................
................................
................................
..............

5

Catching Bees on Flowers with Baggies and Kill Jars

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................................
..............................

5

Bee Vacuum

................................
................................
................................
................................
............................

5

Chlorocresol Humid
or

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................................
.............

8

Pinning 101
:

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............................

9

Labels

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....

11

Pens

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17

Entering Specimen Data

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.......

17

Microscopes

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..........................

18

The Bee Bowl Trap

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20

Glycol Pan or Bowl Traps

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......

27

Flower Traps

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28

Trap Holders

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.........................

29

Field Trip Check
list

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................

30

Bee Monitoring Discussion List and Announcements

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................................
..........................

30

Quick Bee Survey Protocol

................................
................................
................................
................................
...

30

Pr
ocessing Bees that Have Been Stored in Alcohol of Glycol
................................
................................
................

31


Cleaning Bees that Have Gotten Moldy

................................
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................................
................

35

Re
-
hydrating Bees that have Been Pinned

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............

35

Inexpensive, but Powerful LED and Florescent Light Sources

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..............

35

How to Make a Pizza Insect Pinning Box

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36

T. Mitchell’s Guide to the Bees of the Eastern United States

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...............

38

Mike Arduser’s Keys

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.............

39

Canadian Identification Guides

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............................

39

Stylopized Bees

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44

Taking Pictures of Bees with the COOLPIX 990

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................................
....

45

Affixing bee wings to microscope slides

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...............

52

Taxes and Specimens Donations (U.S.)

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.................

53

North American (North of Mexico) Introduced and Alien Bee Species

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................................

53

Mini
-
summary of the
Genera of Eastern North American Bees

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................................
...........

55

Pronunciation Guide to the Bee Genera of the United States and Canada (and Selected Subg
enera)

...............

62

Glossary of Bee Taxonomic Terms

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................................
........................

66

Bee Body Part Figures

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................................
...........

69


W
here to Find Bees


Bees are nearly ubiquitous; they occur essentially everywhere. However, in any given landscape there are
usually a few good places to collect bees where they are
concentrated, diverse, abundant, and easy to capture
and there are many, many places where bees are difficult to find and collect. If you are interested in
biodiversity, and taxonomic surveys, it will be important to discover these hotspots. In North Ame
rica, in
general, good collection locales will be places where floral composition is concentrated or unusual. If you are
unfamiliar with an area then exploring road/stream/river crossings, powerline rights
-
of
-
way, railroad track
rights
-
of
-
way, sand and g
ravel operations, open sandy areas, and wetlands are good places to start, In areas
with a lot of development, the industrial sector often contains weedy lots and roadsides that also can have
good numbers of bees. Note that just because there are few or

no plants blooming (to your eye!), this doesn’t
mean that there are no or no interesting bees present. A good collecting strategy is to put out bee bowl traps
(see sections below) in the morning, and return to good potential collecting sites that you spo
tted that
morning during mid
-
day.

Killing Bees to Study Them

In bee work we almost all are

confronted by the
issue
of having to kill the things we study and explaining that
to the public
as well as

to land managers, Jessica Rykken

pointed out a good essay on that topic at:
http://www.biology.ualberta.ca/bsc/pdf/whywekillbugs.pdf

Nets

Almost any type of insect net will catch bees. However, bee collectors do h
ave preferences. Most people now
use aluminum handled nets rather than wood. Some prefer the flexible strap metal netting hoops, as these
work well when slapping nets against the ground to capture low flying or ground resting bees. Others prefer
the mo
re traditional solid wire hoops. Hoop size varies from about 12” to 18.” The larger the hoop, the
greater the area of capture, however, larger hoops are more difficult to swing quickly due to air resistance and
there is more netting to snag on branches.


BioQuip makes a net which is very portable for travel or backpacking. The pole disconnects into 3 small
sections and the hoop can be folded into itself. Additional sections can be added to reach into out of the way
places. Telescoping poles are al
so available but must be treated with care or their locking mechanisms will
jam. An inexpensive long pole can be rigged by attaching a net hoop to a section of bamboo with hose
clamps. Aerial nets, rather than beating or sweep nets are normally used aro
und the hoops. A fine mesh net
bag rather than the traditional aerial net bag can keep the smallest Perdita from escaping.


Netting Technique

Always hold your net in a “swing
-
ready” position. One hand should be below the head and the other towards
the
back or middle of the pole. Hold the tip of the net lightly against the pole with the hand near the head so
that it does not drag in vegetation. When you start your swing drop the tip of the net.

Bees are best detected by their motion, rather than their s
ize and shape. The mind detects motion much
faster than it can process colors and shapes into bee/not bee categories. Train yourself to key in on
movement; over time you will become more adept at separating bee motion from plant motion.

Bees are lost when you hesitate or check your swing. If you see something that looks like a bee, capture it in
your net. Once in your net you can decide whether or not to keep it. If you spend any significant time thinking
about whether you should or sho
uld not swing, the capture opportunity will be missed as the bee will have
moved on.

Always keep a mental check for the presence of thorny plants in the area where you might swing
-
for obvious
consequences to your net. Additionally, in some areas some p
lants have seeds that can implant themselves
directly into the netting; if that is the case then you might try moving from the usual coarse weave net bag to
the fine weave type that BioQuip sells.

When swinging a net, speed is important as well as follow
-
t
hrough. Bees are very visual and very fast. If you
are timid in your swing or cut your swing short bees will evade the net. Center your net on the bee if at all
possible even if it means having to plow through some vegetation. When a bee is flying low
to the ground it is
better to slap the net over the bee than it is to try to catch it with the corner of your net.

All else being equal, it is better to swing at a bee that is just flying into or away from a flower than a bee that is
actually on a flower.

Particularly if you are trying not to damage the plant, a less than vigorous swing of the
net will simply push a bee on a flower under the net and it will fly away afterwards. After some practice you
can bring your net up to a bee on a flower, wait for
the bee to leave the flower, push the flower out of the way
with your net and still easily capture the bee.

When looking at a clump of flowers that could contain bees stand 4
-
8 feet away. Most people stand too close
to the flowers, which can scare away s
ome of the bees you are interested in, limit both the number of flowers
(and therefore bees) in your field of view, and limit your depth of field. In this way you can view a large area of
flowers, spot a bee, and either lean forward or take one step to pu
t that bee into your net. If you have to take
2 steps or more, you are too far away.

On any flower patch, concentrate on the difficult to obtain bees first. In particular, look for bees that are
moving very quickly, from flower to flower, and try to pred
ict where they will move next.
U
sually there is
some pattern to their flight and often they will come back to the area after making their circuit. Some of
these individuals never really come to rest and you have to swing ahead of where you think you are

going to
catch them. It also pays to look below flower clumps for low
-
flying bees. Some of these are nest parasites,
while others simply prefer to move between clumps of flower just above the ground or grass.

Open soil of any kind and, in particular,

south facing slopes, overturned root masses, clay banks, and piles of
construction dirt or sand should be scanned both for bee nests and for low cruising nest parasites. Nest
parasites (in particular Nomada) usually fly just above the soil in erratic fli
ght paths. The best way to capture
them is to slap the entire head of the net over the bee and quickly lift the net bag up while leaving the rim on
the ground. The bee will fly upwards rather than tring to sneak under the rim. Often this can take several

seconds, so patience should be applied.

There are two ways to catch multiple individuals in a net. One way is to turn your net head sideways after
capturing a bee, allowing the net bag to close over the head and hoping that the bee will not find a way
out.
The other is to physically hold the bag closed above the tip containing the bees (note, in between swinging at
bees, you will be holding the closed net against the pole as you carry it from place to place). In both cases you

will have to periodically

snap the contents of the net to the bottom. Do this vigorously or some wasps (in
particular) may not go to the bottom, and you could end up grabbing them through the net with obvious
consequences to your hand.

In general it is easier to see bees through
the mesh, if you go into the shade or shade the net with your body.
Some people favor green nets over the traditional white ones to reduce this phenomenon.

A video that demonstrates how to use a net to collect bees can be seen at:
http://www.youtube.com/watch?v=n6ZFlz3uA7E

Removing Bees From the Net


Time spent removing bees from the net is time spent not capturing bees; therefore, think about how you are
removing bees from your net to see if you
can speed the process up.

In the beginning, there is usually a great fear of being stung by your subjects. In reality, in North America, only
Polistes, Vespinae, Bombus, Apis, Pompilids, and perhaps a few of the other wasps have significant stings.
Th
ese are large insects and can be readily discriminated. However, even these species do not sting
while
caught
in a net, unless they are physically grabbed or trapped against the net. Thus, over time you should
concentrate on diminishing your fears, and s
pend more time sticking your hand and kill jar directly into the
net. If you are putting your net on the ground to remove bees, you are taking too much time. Kill jars should
be fully charged to quickly kill your specimens, and it helps to have multiple
jars available (see section on kill
jars).

The most efficient means of collecting large numbers of bees is to use vials or containers of soapy water. In
that way you can fill your net with bees and only have to
empty the net
periodically rather than af
ter catching
an individual bee or small number
s

of bees. However, cleaning and processing bees killed
in liquids
requires
some care to do properly (see section on washing and drying bees).

Laurence Packer has gotten to the level where he simply uses his

fingers on all bees except bumblebees; he
gets stung, but says it's all very minor, unless he gets stung repeatedly on the same spot. Aspirators can also
be used to remove minute bees (such as
Perdita
) if you only have traditional killing jars.

Once you
have captured a bee or bees in the net, there are several ways to remove them. In all cases, it is best
to vigorously snap the net to drive the insects to the bottom. You can then safely grab the bag just above
where they are resting. Even the larger an
d more aggressive bees can’t get at the hand that is closing off the
net, due to the bunching of the netting. If you are timid, are worried about the specimen escaping, or have
numerous insects in the net, you can kill, or at least pacify your catch, by
stuffing the specimens and the
netting into your kill jar, closing the lid loosely. Keeping your jars well charged with cyanide or ethyl acetate
will ensure that the specimens quiet down quickly, and you will not waste a lot of time waiting. Once your
specimens are immobilized, you can open up the net and drop them directly into the kill jar without worry.

Most collectors take a more direct approach and bring the open kill jar and its lid into the net, trapping the bee
against the netting. Slapping t
he hand on top of the kill jar through the netting is at times useful to drive the
bee to the bottom of the jar. This can help prevent bees from escaping when you put the cap on. More than
one bee at a time can be put into a bottle this way, but at som
e point, more escape than are captured. Laying
the net frame on the hood of a vehicle at a spot that fits, can help reduce escapes.

Because seeing the bees through the netting can be difficult, (hint: use your body to shade the netting to
better see th
e bees), some collectors have taken to hanging the net on the top of their head. Use one hand to
hold the net out and up, and then use the other hand to reach in and collect the specimen with the kill jar. It is
important in this situation to keep holding

the net out so the bees move away from your head

(duh!)
. Use
small collecting jars, aspirators, or large test tubes that can be handled easily with one hand. Despite having
your hand (and sometimes your head) in the net with the bees, most collectors a
re rarely stung.


In general, bare hands are recommended when removing bees from nets. Bees and wasps will almost never
sting in a net, if you don’t trap them in your hands or against the netting. Use of a centrifuge tube filled with
soapy water makes
removal easy, as you can keep well away from the bees. Some people will use gloves, such
as handball gloves, welder gloves, latex dishwashing gloves (though stinging can occur through latex), and
goatskin beekeeper gloves.

A video that demonstrates how to

remove bees from a net can be seen at:
http://www.youtube.com/watch?v=n6ZFlz3uA7E

General How
-
to Videos on how to work with insect collections are available at:
http://www.bugs.nau.edu/learning_modules.html


Using Ice and Dry Ice

If it is important to keep bees alive or very fresh, bring a cooler of ice or dry ice and two nets. You can
continuously colle
ct bees with one net, and once it is full, place the entire net end into the cooler. If the cooler
is filled with ice, the bees will remain alive but inactive; if the cooler is filled with dry ice, they will freeze. You
can then continue collecting with
a second net. Once that one is full, the bees in the first net have already
been chilled or have perished, and you can transfer them to jars in the cooler for further storage.

Catching Bees on Flowers with Baggies and Kill Jars

These systems are particula
rly useful when working with individual specimens on individual flowers. Pop the
open end of large baggies over flowers with bees on them. The bees can then be sealed in the bag and placed
in a cooler of dry or regular ice for preservation until taken ba
ck to the lab.
Similarly
, putting a kill jar over a
flower and tilting the flower into the jar works to preserve the flower.

Bee Vacuum

The first section on converting a Leaf Blower was contributed by Julianna K. Tuell…

Sam Droege asked me to send out a

detailed description of the modified leafblower that was used in Michigan
to collect flower visitors, because it may be of interest to members of this listserv. Every method used to
collect insects has certain biases, but we found that vacuum sampling end
ed up collecting similar numbers of
both large and small bees to those recorded during timed observations at the same flowering plots by trained
individuals. One obvious advantage of vacuum sampling is that it can be conducted by someone with very little
t
raining.

A Stihl leafblower and vacuum converter kit were purchased from a certified Stihl dealer. My colleague, Anna
Fiedler, who purchased the components and conducted most of the sampling, said it was very easy to
assemble and use. She added two screws

a couple inches from the end of the intake tube (not sure if this was
part of the kit or if this was something extra she did on her own), so that she could use rubber bands to hold a
handmade mesh bag (made of no
-
see
-
um mesh) over the end for collecting t
he insects. She vacuumed each
1m^2 plot's flowers for 30 seconds and then while the leafblower was still on, she would quickly remove the
mesh bag, close it and then place it in a cooler to immobilize the insects in the bag so that they could be
transferre
d to a ziplock bag without losing any individuals. In this way she could reuse the mesh bag for
another sample on the same day and she only needed to carry 4 mesh bags.

Here is the link to the actual model leafblower that was used:
http://www.stihlusa.co
m/blowers/BG55.html


You can find out more details on the natural enemies part of the project via these two references:

Fiedler, A. and Landis, D.A. 2007. Attractiveness of Michigan native plants to arthropod natural enemies and
herbivores. Environmental
Entomology 36: 751
-
765.


Fiedler, A. and Landis, D.A. 2007. Plant characteristics associated with natural enemy abundance at Michigan
native flowering plants. Environmental Entomology 36: 878
-
886.

The manuscript for the bee part of the project has been pu
blished:

Tuell, J.T. and Isaacs, R. (2008) Visitation by wild and managed bees (Hymenoptera: Apoidea) to eastern U.S.
native plants for use in conservation programs. Environmental Entomology 37: 707
-
718.

Another technique for converting a portable “dustbu
ster” vacuum was written by Glenn Hall and is available in
pdf format at our Listserv website (
http://tech.groups.yahoo.com/group/beemonitoring/files/
)

Smaller, Pocket Sized Bee Vacuu
m
-

This section was contributed by Cheryl Fimbel, based on the initial
suggestion by Priya Shihani.




I am writing to pass along a tip that was provided to me by Priya Shihani. She recommended a small hand
-
held
vacuum unit


the Dirt Devil Detailer (mo
del CV 2000) for vacuuming up small bees and other insects off
flowers. This small vacuum does a fantastic job of scooping up all sizes of bee from the smallest to big’uns
(Bombus, but perhaps not the queens). It is especially useful for the tiny bees th
at would get lost in a net, or
are foraging among flower parts that preclude capture with a net. It is ready to use right off the shelf, as it has
a small flap that comes down to prevent escape by insects when the vacuum motor turns off. It is small
enou
gh to fit in a pocket, and one charge of the battery lasts for weeks. I like carrying two of them in a
‘holster’ I devised, like a pair of six
-
guns…. ever at the ready to scoop up a flower visitor, with each vacuum
dedicated to a specific flower species.

It is the most fun I have ever had vacuuming (I just hope my house
guests don’t notice that my flowers are cleaner than my carpet!).

Tracy Zarillo finds the vacuum mentioned in the reference well works well for her.
Osborne, K.H. and W.W.
Allen.

1999. Allen
-
Vac: An Internal Collection Bag Retainer Allows for Snag
-
Free Arthropod Sampling in
Woody Scrub.

Env. Entomol. 28(4): 594
-
596
….specifically refurbishing
a Craftsman Model No. 358.797310
from Sears, Roebuck and Co.

Bees Through Binoculars
-

For those investigators who do observations of bees on flowers or around nest
sites, the Pentax Papilio 8.5X21 binoculars are ideal. They have high magnification and focus down to 0.5m,
permitting sight identifications and detailed behavioral observatio
ns (once you have learned to identify
specimens under the microscope).

Kill Jars

-

Several companies make chemical based kill jars which use either ethyl acetate or potassium cyanide
as the killing agent. There are advantages and disadvantages to both typ
es.

Ethyl acetate

-

Traditional jars are made of glass with a layer of plaster of paris at the bottom. At the start of
the collecting day, pour enough ethyl acetate into the jar so that it soaks into the plaster, but leaves no liquid
on top. If you use
the jar regularly, then the ethyl acetate will need to be recharged every couple of hours, as
it will evaporate. The advantages of using ethyl acetate are: less toxic than potassium cyanide, not a

controlled substance, and relaxes the specimen, which is
useful if the genitalia are being pulled. The
disadvantages are: needs to be replenished often (requiring either that ethyl acetate be brought into the field
or that several charged kill jars remain available), can cause the jar to “sweat” inside which ma
y mat a
specimen’s hairs, significantly degrades DNA
, and will outgas in a hot car
.

Potassium cyanide

-

Most collectors eventually end up using a cyanide
-
based kill jar. BioQuip

makes kill jars
with a hollow plaster top underneath the lid that can be charged with potassium cyanide crystals. However,
cyanide jars can be made from any glass or plastic container. Place a layer of cyanide crystals in the bottom of
the container.
Next add a layer of sawdust. Finally, pour wet plaster of paris over the sawdust. Leave the
jars open for a few hours outside or in a hood, and then close them. Alternatively, a combination of cotton
balls and tightly rolled paper towels can be used in

place of the plaster and sawdust. The advantages of using
potassium cyanide are: knocks down insects quickly, does not significantly degrade DNA, can remain effective
for over a year, and does not add moisture to the jar. The disadvantages are: is ex
tremely toxic, is a
controlled substance, and can change the color of some bees (particularly yellows become orange or reddish),
if bees are left too long in the jar.

Cyanide jars usually work immediately in the field, but if they don’t knock down specimen
s right away, a drop
of water or a bit of spit (don’t lick!) will cause the crystals to begin giving off gas. Many collectors use test
tubes or narrow vials with a cork top as collecting vials. These are useful when there is a need to keep
collections se
parated in the field, such as when collecting off different plant species. Tubes can also be
handled easily with one hand while in the net. Vests, aprons, hip packs, and carpenter belts are useful ways to
keep a number of collecting vials handy.

Most peo
ple will wrap the bottom of glass jars and vials with duct tape to reduce the chance of breakage in a
fall. Additionally, it is handy to place a bit of paper towel in the bottom of each jar to absorb the extra
moisture and regurgitated nectar from the bee
s collected.

After bees have been placed into a well charged kill jar, they usually quiet down in just a few seconds. If the
specimens are taken out of the jar too soon, some may “wake” back up and begin to move again, albeit usually
only very slowly.

Usually thirty minutes or so in the kill jar will prevent this.

Soap Jar

or Tube

-

An alternative to chemical based kill jars are containers filled with soapy water (a mix of
water with dishwashing detergent) or alcohol. These are particularly useful f
or those of you who store
specimens in alcohol, or wash them prior to pinning. The best jars/vials have a tight fitting lid and are large
enough to hold a fair number of bees. They should fit in your pants pocket and be easy to hold in one hand
along wit
h the lid. F
ill the vial about half full with soapy water.

The jar will form a constant head of suds while riding around in your pants pocket. Using it in the net has the
great advantage of immediately trapping any insect in the suds, thus permitting
you to clean out the net of as
many specimens as you wish. With a chemical based (cyanide, ethyl acetate) kill jar, you can accumulate 2
-
4
specimens with some effort, but at some point, more would be leaving than going in. The soapy jar is
particularly n
ice when dealing with large, nasty specimens. The Patuxent lab favors using the large centrifuge
tubes, as they slip into the pocket easily.

You have to be a bit more aware of how you carry the jar when open (water seeking its own level and all that),
b
ut such jars can also easily be used to directly collect off of flowers without a net.

Specimens can be readily left in the soapy water for 24
-
hours and, while a bit soggy, will even last for 48
-
hours
without too much degradation. Afterwards, specimens
can be either dried and pinned, drained and put into
alcohol for long
-
term storage, or drained, wrapped with a piece of cloth (to soak up excess moisture and to
prevent breakage) and frozen in a plastic bag. Specimens look best if cleaned and dried withi
n 24 hours of
capture in bowls or soapy water, if cleaned immediately after capture some specimens can “wake
-
up.”
However, this can readily be checked by freezing any specimens that do begin moving.

The advantages of the Soap Jar are:




Don't have to lug t
oxic chemicals around



Soap and water are readily available



Restrains specimens immediately



Can collect all specimens in a net at one time



Inconspicuous to the general public



Pollen and gunk are washed off while in the vial



Cheap

Disadvantages:



No pollen
analysis



Specimens are wet



Jar needs to be held a bit more upright when open than a normal killing jar



If cap not on correctly, the water can leak



Specimens have to be dried prior to pinning

Chlorocresol Humidor


(Contributed by Rob Jean)
-

For those of us

that enjoy net collecting, but do not have the time to prepare and
pin up our day' s catch the same evening, here is a technique for preserving specimens in a pliable state for
extended periods of time (6 months
-
1yr or longer if moisture conditions are ke
pt right). This is a simple
technique I learned from Mike Arduser, Natural History Biologist, Missouri Department of Conservation, who
uses it exclusively and rarely pins anything until he runs it through a chlorocresol humidor. The technique
requires: 1
.) a pint or quart
-
sized plastic container with a tight seal (I use a 4 cup or 1 qt Ziploc Twist N Seal
container, but I have used on occasion up to 1/2 gallon sized containers) 2.) paper towels, 3.) chlorocresol (an
antifungal crystalline substance with

a sugar
-
like consistency available from Bioquip
-
item #1182B
-

$18.45/100
grams) (chemically = p
-
chloro
-
m
-
cresol or 4
-
chloro
-
3
-
methylphenol), 4.) a few strips of duct tape or its
equivalent, and 5) a few drops of water.

To make the humidor, start by puttin
g one rounded teaspoon of chlorocresol in the middle of one heavy paper
towel or two lightweight paper towels. Then fold the paper towel(s) around the chlorocresol so that the
chlorocresol is enclosed in the paper towel(s), and so that the folded paper to
wel(s) can fit into the bottom of
the plastic container. Tape the loose edges of the paper towel(s) with narrow strips of heavy (duct) tape, using
as little tape as possible. Thus, the container will have a securely sealed, but porous, chlorocresol "packe
t" at
the bottom.. You should do this under a fume hood or outdoors as chlorocresol has a strong smell and it can
be harmful if inhaled or swallowed.

Once the chlorocresol packet is in the container, you simply have to play with the moisture level to get i
t
perfect. In most cases, keeping the paper towel damp (not soaked) is enough to keep the specimens moist
and pliable enough to spread mandibles and pull genitalia, sternites, etc., but you will probably have to
experiment a bit with this before you get i
t right. Specimens will dry up and become brittle if there is not
enough moisture (but can be rehydrated in a few days usually). If there is too much moisture, hairs will
become matted on specimens and make them harder to identify later. Again, you may h
ave to play around
with the exact moisture conditions for the container/humidor you are using. One good thing is that the
chlorocresol goes a long way (10 years or longer according to Mike A.). I have been using this method for two
years and I am still o
n my original doses of chlorocresol in my humidors (I carry two with me at all times when
collecting).

After I have the humidor, I can catch specimens on flowers without an immediate need to pin. I can keep each
collection event (different flower species, times of the day, etc.) in separate glassine envelopes or paper
triangles within the humidor. Glassin
e envelopes and paper triangles are great to use in this situation because
they are easy to write data on, and because they allow the moisture in the humidor to get to the specimens.
With periodical checking on the moisture levels in the container (I have

to check mine every week or two),
specimens can last several months to a year according to Mike Arduser. The specimens stay fresh as the
chlorocresol wards off fungal agents. The chlorocresol also seems to relax specimens somehow, which makes

mandible s
preading and genitalia pulling a little easier in bees.

One caution: pollen loads (particularly apines and panurgines) can become soupy in the humidor and may
inadvertently get stuck or plastered onto other bees. Also, specimens will smell like chlorocre
sol for some time
after they come out of the humidor. Good luck and I hope this method saves some preparation time.

Pinning 101
:

Types of Insect Pins to Use

-

Bees are usually pinned using pin sizes 1
-
3, with size 2 being the most common.
Pin size 1 is p
rone to bending when pressed into traditional hardboard lined trays and boxes, but does nicely in
foam units. Pin sizes below 1 should not be used as they are delicate, do not hold labels well, and end up
bending if the specimen is moved or viewed often.

Size 4 is generally too large for anything other than
bumblebees. In humid environments, stainless steel pins should be used to prevent rusting. Student pins
should be avoided as they are cheaply made; the tips bend and the balls come off. Insect pins
can be
expensive. The cheapest way to purchase them is to order in bulk directly from Czechoslovakia, where
apparently most are made. Some newer inexpensive (same price as European steel pins) stainless steel pins
are now available from China that appear

to be of high quality.

Traditional Pinning

Techniques

-

Bees can be pinned directly from the killing jar into boxes, or they can be
washed first. If the bees are dry and not matted down, then pinning directly to a collecting box is best, as it
preserve
s the pollen load for future analysis and speeds up the entire process. However, if the bees are
matted from too much moisture and regurgitate, wash and dry them using the protocols listed in this manual.
They will result in better looking, easier to ide
ntify specimens. If the pollen load is not going to be analyzed,
then washing the specimens also has the advantage of eliminating the pollen from the scopal hairs and
diminishing the “dustiness” of the specimens.

Each person develops his or her own proc
ess when pinning bees. Some pin under the microscope, which
usually results in very accurate placement of the pin, but many pin by eye. One technique is to hold larger
specimens between the thumb and forefinger with the pin ready in the other hand. Use

another finger from
the hand holding the pin to help hold the specimen steady while inserting the pin accurately into the bee’s
scutum.

Others pin larger bees using a pair of forceps or tweezers, trapping the specimen on a foam pad. Expanded
polyethyle
ne foam (often referred to as Ethafoam) or cross
-
linked polyethylene foam (our preferred foam) is
better than polystyrene foam (usually referred to as Styrofoam) for pinning purposes. Styrofoam is not
supportive enough; both labels and specimens will bend

too much when pinned upon Styrofoam.

Specimens are best pinned through the scutum between the tegula and the mid
-
line. The midline of the
scutum often contains characters that are very useful in identification, which can be destroyed by a pin. Most
m
useums prefer that specimens be pinned on the right side.

For someone new to pinning, use of a purchased insect pinning block is helpful to determine the correct height
a specimen should be placed. With experience, one can use pieces of foam of the co
rrect depth, or even
adjust specimen height by eye, which will be the quickest. Remember to leave enough room at the top of the
pin so that the specimen can be safely picked up by the largest of fingers. Equally important, leave enough
room at the bottom

for two or more labels and room for the pin to go into the foam of a collection box.

A video that demonstrates how to pin bees can be seen at:
http://www.youtube.com/user/swdroege#p/a/u/1/V2F8LBQV5L0

Gluing Small Specimens

-

If specimens are too small to be pinned, they can be placed on a point, glued to the
side of a pin, or attached as minuten double mounts. Reversible glues, such as Elmer’s G
lue Gel, white glues,
tacky glue, clear nail polish, shellac, hide glue, and others should be used.

Gluing to points: The use of points is traditional. Points are very small, acute triangles cut from stiff paper

using a special punch, which can be orde
red from entomological supply houses. Place the pin through the
base of the point. Elevate the point on the pin to the same height as a pinned specimen. Glue the small bee to
the tip of the point, usually on its underside.

Gluing directly to pins: Wh
en gluing a specimen directly to a pin, rather than to a point, the specimen is glued
on its side or the underside between the thorax and abdomen. Again, most museums prefer that specimens
be glued on the right side. Gluing specimens to the side of the pi
n has the advantage of speed, better
prevention of glue hiding useful characters, and a specimen that is easier to view under the microscope. Its
axis of rotation is minimized and the point is no longer there to hide the view or block the light. Specimen
s
should be glued to the pin at the same height as those that are traditionally pinned.

In the past, we have used white, tacky glue in our lab. This is a thick glue which sets up within seconds. It
allows the glued specimen to be set upright in a box

immediately, without the danger of it losing its placement
on the pin. From our limited investigations, Aleene’s Original Tacky Glue in the gold bottle or archival paper
glue appears to be the best gripping, tacky glue.

We now like to use glue gels w
hen pinning bees. Glue gels have a longer work time, dry crystal clear and are
easily reversible. Because the set
-
up time is longer than tacky glue, leave the pin resting on the specimen for
at least 5
-
10 minutes prior to picking it up. Parchment paper
is very helpful to have around when gluing bees.
It is a silicone impregnated piece of paper that can withstand the heat of an oven but is super slick. It provides
a “non
-
stick, Teflon
-
like” substrate on which to work, because glue does not adhere well t
o it. Another nice
thing about parchment paper is that dried specimens can be easily positioned on it. They will slide around
without sticking or breaking. We now dump dried specimens onto the paper and pull up the sides, which
causes the specimens to s
lide into the center. Once in the center, they are positioned in a line which makes
pinning even more rapid. At this point, you can pin the paper to the top of a large foam board. Place pins
with glue at the proper height onto specimens. After the glu
e is set, press the pointed tip of the pin with your
finger. This will cause the specimen to rise up, allowing you to grasp the top of the pin and move it into a
collection box.

A video that demonstrates how to glue a bee to a pin can be seen at:
http://www.youtube.com/user/swdroege#p/a/u/0/9KfLCmYOKtA


Current BIML techniques:

While unorthodox, our current process for pinning involves: washing and drying
specimens in the mac
hines listed in this document, placing them in open, labeled Petri dishes and letting them
completely dry for a week or more prior to pinning. If time doesn’t permit pinning right away, after a week or
so of drying, the Petri cover is replaced and taped on
, and the specimens are stored in their dishes.

When ready to pin, all the specimens are laid out on a large foam pinning board covered with parchment
paper and a pin is glued to the side or underside of each ….including the largest specimens…. using ge
l glues.
Large specimens require larger amounts of glue, and all specimens need to have pin and glue attached to the
body of the specimen rather than to a wing or leg. We use a magnetic pin holder that attaches to the wrist.
These are available in hardwa
re stores, online, or in sewing shops. A sawn off section of bolt (we use 2 of
them) is handy to have on the wrist holder, as the threads will separate the pins for easier pick up. We then
run a small line of glue on the side of our thumb or index finger

(Thank you Harold Ikerd for this idea) on the
same hand that has the wrist holder.

A reverse set of tweezers is used to pick up a pin by the head or the tip. It is dipped into the glue line on the
thumb at the proper specimen height, and then placed on

the specimen on the pinning board. Because the
specimens are so dry, care must be taken to place the pin gently. The pinned specimen is left on the pinning
board until the glue sets. With a little practice, it is easy to achieve pinning rates of 250+ p
er hour. None of
these gizmos are necessary to glue bees quickly; fingers work nicely without tweezers, glue can be spread
directly from the bottle, and pins are very convenient if stuck into the foam.

After the bees have dried for a couple of hours, they

are then transferred to boxes. In some instances, that
transfer can be accomplished efficiently with the attachment of labels, saving another step. Jane Whitaker has

found that magnetizing her tweezers helps in picking up glued specimens on pins.

Minute
n double mounts:

Minuten double mounts are not used very often, but do create the best looking
mounts. A tiny bit of crosslinked polyethylene foam is pinned to the same height as a regular specimen. A
minuten pin is added to the right side of the specim
en and then inserted into the foam block. On the down
side, this takes a lot of time to accomplish.

General Videos on how to mount and work with insect collections are available at:
http://www
.bugs.nau.edu/learning_modules.html

Bee Boxes

-

There are a variety of drawers, cabinets, and boxes available to hold specimens. We prefer to use
the simple cardboard specimen box with a completely detachable lid, and an Ethafoam bottom for everything,
ex
cept for housing our synoptic collections. These boxes are stackable, the date and location can be written
on the outside in pencil and then erased when reused, are relatively inexpensive, and, unlike hinged lid boxes,
are convenient to use in cramped spa
ces on a desk or worktable. Such boxes can be made from scratch.
Instructions for making “pizza” insect pinning boxes can be found in this document.

After a batch of specimens is washed, dried and pinned, we place them in a cardboard specimen box. At

the
upper left hand corner of the box, a tag with the date, place, site or batch number is pinned. This tag is usually
the original tag that was placed in a batch of specimens when first captured. Pin a line of specimens to the
right of the tag, and con
tinue running from top to bottom, and left to right, like a book, until complete. The
next tag is placed immediately thereafter and so forth until the box is filled. In general, it helps if each box
contains specimens from only one region. We label the y
ear across the top of the box, then the month, and
then the locality, so that we can quickly pick out the box we want.

Alana Taylor
-
Pindar has alerted us to an inexpensive source for quality cabinets and drawers for your
collection at:
http://www.quebecinsectes.com/pages/pages_english/macrodontia_english.html

in addition to
the
http://www.bioquip.com/.

Control of Pests



Simple cardboard boxes are not pest proof. Dermestid beetles are the primary pest of
insect collections. Fortunately, infestations are usually small, perhaps seeing one beetle larvae in a box
scattered here and there. An infected specimen is usually
easy to spot, as small black droppings and shed skin
are visible below the specimen. Control and prevention take place, according to the literature, by freezing the
box at
-
20C (about zero degrees Fahrenheit) for 3 days, thawing for a day and then freezin
g for another 3. In a
pinch, kitchen freezers appear to work too. Mothballs and pest strips can be effective, but carry some
apparent health risks with long
-
term exposure. Spring is a good time to freeze your entire collection, as that is
when dermestid
s appear to be most active. An excellent means of keeping your collection pest free
(particularly if using cardboard boxes) is to keep each box in a large zip lock bag. Note that you should have let
the specimens dry out thoroughly after pinning (one mon
th or so) before enclosing them in the bag.

In humid conditions (such as July and August in Maryland), unprotected specimens, particularly those just
caught, can turn into balls of mold. Either take them into an air
-
conditioned space or put them in plasti
c bags
or tightly closed bins that contain active desiccants. Keeping specimens in a refrigerator or cooler without
moisture control will ultimately lead to mold too.

Labels

Following pinning, labels are produced for each batch of specimens. We use a label generating program
available on the
www.discoverlife.org

web site. Each batch or site is given a unique site number and each
s
pecimen is given a unique specimen number. On each label, the specimen number and site number are
listed, as well as the country, state, county, latitude, longitude, date of collection, and collector. A small data
matrix is present on the label that enco
des the specimen number and permits the specimen to be scanned
with a hand
-
held scanner directly to a database while remaining in the box. These data matrices are included
automatically in the free Discoverlife system (http://www.discoverlife.org/label/)
or can be added using
commercial software such as BarTender (http://www.seagullscientific.com/).
Many a beginning student of

bees has rued the day that they did not give their specimens unique numbers.

Dan Kjar has generalized the Discoverlife label prog
ram so it will print out on a laser printer.
You can use his
simple web based form (
http://bio2.elmira.edu/fieldbio/
)
by following the link at the bottom of the page for
insect labels. Each label is unique

based on the specimen number.

Depending on how many labels you are making and your Internet speed, it will take a little time to build the
label page. 50 labels take about
1

minute

to assemble. The system will be integrated into Discoverlife soon
and whe
n that occurs Dan will announce that on his site.

In a good museum cabinet, specimens deteriorate only very slowly and can last for well over 100 years. That is
not true of the paper used in making labels. Paper that is not archival or acid free gradua
lly deteriorates.
Fortunately, archival paper is readily available in office supply stores. A heavier weight paper is also important
to use so that the label stands up to handling and the pinning process. A 35
-

65 pound paper is good label
stock.

Spec
imen labels are quickly added to specimen pins by laying them across a piece of Ethafoam
-

the thickness
the desired height of the label on the pin. To increase the durability of the Ethafoam, glue it to a piece of
plywood, which will form a sturdy pinnin
g surface. To manufacture a pinning board, smear white or wood glue
across both surfaces, rub together, and then place another (unglued) board on top of the foam. Pile books or
other heavy objects on that board to clamp the foam and board tightly togethe
r. Let dry overnight. It can
then be used as is, or the edges can be trimmed with a saw for a nice and tidy look. Labels are oriented along
the same axis as the specimen. Prior to putting labels on specimens, do a quick check to make sure the label
in
formation matches the row tag.

Cutting out labels can be a time consuming aspect of any project. We speed up the process by cutting out
rows of labels;
placing them in their box and then cutting the individual labels apart with scissors. See:

http://www.slideshare.net/sdroege/preparing
-
insect
-
labels
-
a
-
faster
-
way

and
http://youtu.be/HqxrkC6xe40
.

Ray Geroff

uses a surgical/dissection scalpel and handle. He prefers the #4 handle with a #21 or #22 blade. It
works well for cutting the strips but works really well when cutting the individual labels apart once they are in
single strips.

Gretchen LeBuhn has a
system for making labels in Word, which is explained below.

Open up a new Word document and just type the label as you want to see

it, i.e.,


CALIFORNIA: Napa Co.

Rector Reservoir, 60m

3.2 km NE Yountville

38º26'13"N,122º20'57"W

17 March 2002, ex: Vi
cia sativa

G.LeBuhn, R.Brooks #2002001


As a numbering system, make the bees collected at a single species of plant an individual collection record.
For example, bees collected on
Vicia sativa

at Rector Dam are collection #1 and those collected on
Lupin
us
bicolor

are collection # 2. Keep this system going or some similar system so that you can identify and talk
about each collection separately each year. You can use #2002001 for this year, and then start over next year
with collection #2003001, etc. Th
e point is to adopt some system by which you can talk about any particular
collection event in a multi
-
year study and that it has a numerical identifier.


Now back to making labels…


I make a label log which I actually type directly into my data base and
then extract and put into Word. I cut and
paste a copy of each collection event the number of times needed to label the bees in each lot. I do this in one

long continuous roll. When I am finished, I put it into column format to fit more per page.


Now I h
ave all of my labels duplicated like this:


CALIFORNIA: Napa Co.

Rector Reservoir, 60m

3.2 km NE Yountville

38º26'13"N,122º20'57"W

17 March 2002, ex: Vicia sativa

G.LeBuhn, R.Brooks #2002001

CALIFORNIA: Napa Co.

Rector Reservoir, 60m

3.2 km NE You
ntville

38º26'13"N,122º20'57"W

17 March 2002, ex: Vicia sativa

G.LeBuhn, R.Brooks #2002001

CALIFORNIA: Napa Co.

Rector Reservoir, 60m

3.2 km NE Yountville

38º26'13"N,122º20'57"W

17 March 2002, ex: Vicia sativa

G.LeBuhn, R.Brooks #2002001

The abov
e was for 3 bees collected in Collection #1. Leave a blank line between collection events to see
where each collection event starts.


3) Click "Edit"… select "select all".


Click "Format"… select "Font"… type into the "Size" window the number 3 (

for 3 point font) and click okay.


Click "Format"… select "Paragraph"… select under "Line Spacing" the word "Exactly"… under "At", select "3
pt." (this sets the leading or space between lines)


Click "Format" … select "Columns"… under "Number of Colum
ns" start with 8… under "Width and Spacing"
set the "Space" (that is space between columns) to 0.00. Check with Print Preview, which is selected after
pulling down the "File" menu. The trick here is to get the columns as close as possible to each other
without
any lines wrapping around. Sometimes I can get 9 columns, and other times when the label lines are longer I
can only get 7 columns. 8 columns is my usual maximum column width.


You are done, and can now print onto your acid free or archival, 100%

linen ledger #36 white paper. Cut the
labels out neatly, not leaving white around the edges, and place the labels on the specimens with the top of
the label on the right with the specimen's head going away from you.

Hannah Gaines uses Microsoft Word’s Mai
l Merge to Efficiently Create Labels…

Directions for making specimen labels using Mail Merge in Microsoft Word

First you need to have all of the information you want to use in an Excel spreadsheet. We usually don’t use
the master database for this. Inste
ad, save a copy of the master database (ex
“NSF2006_specimen_database_6_14_LABELS”). You should have a separate column for each piece of
information you need on the label (state, county, coordinates


decimal degrees or lat/long, site name, date,
collecto
r, unique ID).

Sample label:

USA NJ Mercer Co.


40
o
19’9”
-
75
o
22’29” Starkey

15 April 2006 K. Ullmann

ID # 106


Actual size:

USA NJ Mercer Co.

40
o
19’9”
-
75
o
22’29” Starkey

15 April 2006 K. Ullmann

ID # 106


(This is just a sample for formatting purposes and
is not accurate at all.)

Now open a blank Word document.

Go to “Tools”, “Letters and Mailings”, “Mail Merge Wizard”. A wizard panel will show up on the right of the
screen.

Select document type “Directory”. Click “Next: Starting document” at the bottom
of the panel.

Choose to “Use the current document”. Click “Next: Select recipients”

Select “Use an existing list” and then click “Browse”

Select your “…_LABELS” document.

Select the correct sheet (ex “DATA”)

Choose “Select All”, “OK”

Click “Next: Arrange
your directory”

Now you will use the “Insert Merge Field” to arrange the labels (note that you can also
type words

that will be
in every label so you don’t need a separate field in your database)

Sample layout (make sure to put two “ENTER”s after your last

field) :

USA NJ

«County»
Co.

«Lat» «Long» «Site»

«Date» «Collector»

ID #

«uniqueID»


Click “Next: Preview your directory”

If the preview looks right, click “Next: Complete the merge”

Click “To New Document”

Before merging the entire database, try just merg
ing records 1 through 20 to make sure it looks correct. If you
are not happy with the way the labels look, click on “Previous: Preview your document” and rearrange it to
make it work for you. When you are happy with it, click “To New Document” again and
select “All” records.

In your new document, select all (CTRL+A) and set the font to “Arial Narrow” size 4. Now click “Format”,
“Columns” and make 12 columns with spacing of 0”. Next select “File”, “Page Setup”. Type in “0” for all
margins (top, bottom,
left, right). Word will auto
-
correct to the minimum margin size to fit the printer. These
settings should be the most efficient in order to reduce paper use.

Before printing, check the bottom of the columns of labels on each page and make sure that your
labels are
not split up across columns. If they are, just add an “ENTER” to put them back together. Now print on normal
paper. Check the document for any mistakes and fix them. Now you can print on the fancy
-
schmancy label
paper (slightly heavier, acid
-
free paper


should be in the file folders on my desk in a folder called “Label
Paper”.

If you have problems with the number formatting not carrying over correctly into Word, take a look at this
website:
http://office.microsoft.com/en
-
us/word/HA011164951033.aspx#field%20codes

These directions seemed to work all right:

Format currency and other numbers by using field codes

Let's start with an example. Say you i
nsert a Price field into a form letter that you're preparing
for a mail merge. In the main document, it looks something like this, where «Price» is the
field:

The gizmo you ordered will cost «Price».

Press ALT+F9, and you'll see the code behind the field.
That code will look like this:

The gizmo you ordered will cost { MERGEFIELD "Price" }.

You can control the formatting of the prices in that field just by typing a few additional
characters (that is, by adding a formatting switch) inside the braces.

To
include:

a dollar sign

four digits by default, and a space if the number you're merging has only three digits

two decimal places

and a comma between the first and second numbers

this is what you type (shown in bold) in the field code:


{ MERGEFIELD "Price"
\
# $#,###.00

}

When you finish typing, press ALT+F9 to stop looking at field codes. Now when you merge,
all of your prices will be formatted exactly the way you want. (You can use this same
approach with numbers other than prices.)

Here's a breakdown of th
e elements in the switch we just used:


The name of the field that you inserted into your main document. It corresponds to a column in your Excel
worksheet.

Backslash, which starts the formatting switch.

Definition of the switch


in this case, to format
numbers.

Characters that you want to include


for example, a
$

that appears before each price.

The maximum number of digits. If there are fewer digits in a number, Word leaves a blank. Type commas
where you want them to appear in the number.

Decimal point
, which you type where you want it to appear. The zeros specify the maximum number of digits
after the decimal point. If there are fewer digits, Word puts in a zero.

In the See Also box, you will find a link (called Numeric Picture field switch) to more
in
formation about formatting numbers by using a switch.

Format dates by using field codes

You can also use a formatting switch to get dates from a Date column in your spreadsheet to
look exactly the way you want in your merged documents. If you insert a Date

field into the
main document and then press ALT+F9, you see this:

{ MERGEFIELD "Date" }

To get all the dates in the merged documents to have the format February 18, 2008
(regardless of how the dates are formatted in the worksheet cells), you can add this
formatting
switch (shown in bold) to the field code:

{ MERGEFIELD "Date"
\
@ "MMMM d, yyyy"

}

In the See Also box, you can find a link (called Date
-
Time Picture field switch) to more
information about formatting dates by using a switch.

Determination Labels



These labels are used to write the species name along with the person who did the

identification (the determiner).
You can email Sam Droege (
sdroege@usgs.gov
) for Excel spreadsheets that
will print out blank dete
rmination labels that you can modify with your name and date.

Pens

When writing locality or determination labels by hand, archival ink should be used. Rapidographs were most
commonly used in the past, but they have almost entirely been replaced by certain

technical pens, as
Rapidographs tend to clog when left unused for any length of time. Technical pens in sizes 05 and 001 are the
best and are available in art supply stores and from entomological supply stores. Be sure that they state that
they are usin
g archival ink.

Organizing Specimens for Identification

After the specimens are labeled and those labels checked against the original row labels in the box, the
specimens can be freely moved about for identification. We usually sort and identify only those specimens in
a single box rather than try to merge spe
cimens across many boxes. Others color code their projects with
colored pieces of paper placed under the locality label, so that projects can be tracked visually in large groups
of specimens. In this way, multiple projects in multiple states of completio
n can be tracked and are less likely
to become entangled.

When identifying specimens, we make a first pass through the box without using a guide. These are taken out
of the box and pinned to a separate foam board. This board is set at a 45 degree angle

next to our
microscope. As new species are detected, a determination label is created (available as a modifiable Excel file
from us). The determination label is pinned to the board separately from the specimens, so that it can be
easily viewed. All su
bsequent specimens of that species are then placed to the right of the label. Bees that
cannot be immediately identified are kept separate and identified at the end using computer and paper
guides.

Once bees are all identified, they are placed back into

their original box. Bees are placed in the box in rows
starting at the upper left corner, and going from left to right, top to bottom with determination labels
interspersed at the beginning of a new group of species. Females are placed so their label is

positioned
vertically and males positioned so that their labels are horizontal. Positioning the sexes this way permits those
who enter the data to quickly ascertain and check the sex without having to check the label.

Entering Specimen Data

In the syst
em that we use, each specimen has a scannable matrix on its label. Data entry consists of scanning
each specimen directly from the box into an Access database. The scanner has a feature that sends a linefeed
character at the end of scanning in the number
, thus moving the cursor down one line to the next cell where
the next specimen can be scanned…and so forth until that species is completely entered. Access has a nice
feature that permits default values for database fields. Thus, genus and species field

defaults can be set to the
current species being processed, and as the scanner enters a number and drops down a line, the data for the
other fields are automatically entered. Data entry becomes simply a matter of pulling the scanner trigger and
periodic
ally resetting species and sex information either by hand or by changing the defaults. Access has
another nice feature which sets off an alarm or sound if a number is entered twice
-

something that can easily
happen in a crowded box of specimens.

After th
e data are entered by one person, another person cross
-
checks the specimens and the database. After
that final check, the bees are dispersed to final resting spots in museums, sent to other colleagues, or their
pins are recycled for reuse.

Shipping Pinn
ed Specimens

T
he box you ship th
em in should have the following

characteristics: The specimens should be firmly pinned
into the foam. Cut a piece of cardboard that will fit snuggly inside of the box and rest on top of the specimens.

(Do not use foam for
this layer as it can engulf the tops of the pins and cause problems when removed.) Place
either pinned specimens or empty pins in all four corners of the box to support the cardboard. Some people
will also pin loose cotton wadding in the corners of the b
ox so that if a specimen comes loose, it will be
trapped by the cotton. Two pieces of tape can be affixed to the top of the cardboard in such a way as to form
handles that will help remove the cardboard without upsetting the specimens below. Simply pre
ss one end of
the tape to the cardboard and then fold the other end back on itself so the sticky sides meet. If there is space
between the top of the cardboard and the lid of the box, put in some bubble wrap or packing peanuts, so that
when the lid is clo
sed it slightly compresses the cardboard to the tops of the pins keeping them in place during
travel. Tape the lid of the box closed. Put the box of specimens into a larger box with at least 2 inches of free
space on all sides. Fill the box with packing

peanuts, bubble wrap, etc. and ship. In the U.S. we have found
parcel post to work fine, albeit not as fast as Fed Ex or UPS. For valuable specimens all companies provide
tracking and confirmation of receipt services.

Microscope
s

When using bowls or ne
ts, it is easy to quickly amass a large collection of bee specimens. Unfortunately,
unlike most butterflies, bees (even the bumblebees) need to be viewed under a stereo or dissecting
microscope to see the small features that differentiate among the specie
s. While even inexpensive
microscopes and lights can be of some use, in the long run they lead to frustration. Inexpensive microscopes
usually have poor optics, very low power, small fields of view, difficult to set or fixed heights, and their stands
a
re usually lightweight and often designed in such a way that makes specimens difficult to manipulate.

Unfortunately, a good microscope is not cheap. New, our experience is that an adequate microscope costs
over $1000, and good ones run over $2000. Tha
t said, microscopes with even moderate care can be seen as a
one time investment. Additionally, because a good microscope has optics that can be adjusted and cleaned
(unlike most inexpensive ones), it is usually safe to buy a used or reconditioned microsc
ope from an online
dealer (buying off of E
-
Bay or Craig’s List is more risky as the seller has less of a reputation to risk). There are
many used microscope sites; we have purchased microscopes from several of them, and have never had a bad
experience. I
n two cases, the purchased microscopes had a problem, and in both cases, they were repaired for
free. Usually, used prices are about half the cost of new.

Good stereoscope brands to consider that we have experience with include Leica, Zeiss, Olympus, Wi
ld, Wild
-
Heerbrug, Nikon, and Meiji. We can supply you with some model numbers from our collection, or you can
send us web sites with the microscopes you are considering. We will be glad to give you our impressions. Of
special consideration are the Baus
ch and Lomb StereoZoom series. These microscopes have been around for
years, and often form the core of college biology and entomology department teaching labs. These are
adequate to good scopes and we have about 5 in our lab. They are readily available

used from $500
-
$900
online. Their negatives include a view that is not as good as the better scopes and the zoom magnification is
on the top, rather than on the side. Finally, be aware that many of these scopes only go up to 30X power with
the standar
d 10X oculars, though higher powered models exist and higher power replacement oculars are
readily available.

What follows is a list of Microscopes recommended by other Bee Researchers and amateurs.
They range from
high end to low in no particular order.

Zeiss Stemi
DV4
-

about $2000

Leica EZ 4
-

$1150 to $820

(several people responded that they use this line)

Omano Stereoscope OM9949
-

<$1,000

Bausch and Lomb Stereo Zoom 5
-

$150 used

(these are the standard college student scopes of the past)

Leica
2000
-

$850

Leica S6E
-

$1100

Leica S8 APO
-

$3400

Wild M8
-

$1500 used

Wild M3Z
-

$1500 used
-


Olympus SZX12


Olympus SZ60 zoom


Olympus SZ61 with an after market ring
-
light. $2,000
-
$2,400 range

Meiji EMZ
-
5TR body
-

$2000 10 years ago

Leica MZ12.5
-

$6,0
00
-

8,000

Olympus SZX16
-

$6,000
-

8,000

Magnification

-

Magnification power needs some mention here. Any adequate to good scope will have
variable power settings. We have never seen any instance where the lowest magnification was an issue, but a
useful

scope should go up to about 60X power or higher, something that many good scopes do not achieve
with the standard 10X ocular. If the scope does not go to that high a power, it is a simple matter to change the
magnification by purchasing a higher power se
t of ocular pieces (these are the eyepieces that you look into).
Oculars simply slide into tubes on top of the scope and are readily removed (as some of you who have turned
a microscope upside down have found out). However, sometimes there is a set screw
that needs to be
released first. That said, replacement oculars, while almost always available for every model and brand, can
be expensive to purchase. Magnification is determined by multiplying the magnification of the ocular lens
(this number is list
ed usually on the side of each ocular piece, but sometimes is found on the top, and is most
commonly 10X) by the zoom or magnification level which is listed on the zoom knob. Note that some
manufacturers list the zoom levels multiplied out with the assum
ption that you are using 10X oculars.

Most higher
-
end microscopes come with a zoom magnification where all powers are available in any
increment. In some scopes, powers are available only in steps. I haven’t found the scopes that move in
increments to
be any major hindrance. I have found, however, that scopes that have the magnification/zoom
feature available on the sides of the scope in the form of a small knob are the easiest and quickest ones to use.
The ones with the knob on top or located as a mo
vable ring around the base of the scope head take more time
to change. Often the magnification is changed several times when viewing a specimen.

Measuring Reticule

-

Some microscopes come with a measuring reticule in one of the oculars, but most do
not.

A measuring reticule is a very small ruler etched into a piece of glass. These are useful for taking precise
measurements or, more often the case, taking relative measurements. This piece of glass is inserted into the
bottom side of one ocular. All
or almost all oculars are built in a way that they can be taken apart for cleaning.
Often there is a threaded tube inside the body of the ocular that holds the lenses in place. If taking one apart,
be gentle as the threads can be delicate. Measuring re
ticules can be ordered online, or some microscope
dealers will custom make one for you.

Adjusting, Cleaning, and Storing Microscopes



Most good scopes are fairly sturdy and don’t go out of
adjustment without suffering some sort of blow. In our experien
ce, we have come across two primary
adjustment issues: the oculars don’t focus in the same plane, or the images the oculars are processing are out
of alignment. If the images do not completely align no matter how much you play with the width of
adjustmen
t of the eyepieces, the scope probably has significant problems and will have to be repaired
professionally.

Differential focus is usually something you can fix. Small differences in the focal distance of the oculars can be
accommodated by your eyes, but at some point, the eyestrain will become apparent and uncomfortable. In
most scopes, one or both of the tube
s that the oculars slide into are adjustable. These focusing eyepieces are
easy to determine as there are zero, plus, minus, and tick marks to align. To adjust the focus so that both
eyepieces are in the same focal plane, place a piece of graph paper or
something similar on the base of the
scope and shine a good light on it. Adjust eyepieces to zero. If there is one eyepiece that is fixed, then open
that eye and close the other. Change the focus of the microscope so that the grid is in sharp focus. No
w close
that eye and open the other. If the grid is not in alignment, then adjust the focus of that eyepiece until it is. If,
as it sometimes rarely happens, after adjusting in both directions, you still cannot get the eyepiece in focus, try
sliding the
eyepiece up slightly. If that doesn’t work, it is likely the other eyepiece is the one that has to be
adjusted upwards. If the microscope has set screws, you can use them to fix the height; if not, you will have to
work out some other mechanical means.
Usually, however, such an extreme situation indicates that
something is generally wrong with the scope or the oculars. You might check the oculars to see if a lens is
loose or if you have mismatched oculars from some other scope.


The objective lens of a
microscope almost never needs to be cleaned. However, the top lenses of the oculars
often do, particularly if the person using the scope likes to press his or her eyes close and wears make
-
up
(cheap mascara is the worst). We use lens paper and window cle
aner as needed. When the scope is not in
use, put a microscope cover or a large baggie over both ocular lenses to keep the dust out.

Charlie Guevara reports a clever way to modify i
-
Phones into a field microscope at:

http://hacknmod.com/hack/turn
-
an
-
iph
one
-
into
-
a
-
microscope
-
for
-
10/

Holding Specimens and General Microscope Setup

-

Most people when viewing specimens under the
microscope, place them on a piece of clay, foam, cork, or some sort of stand. We try to avoid this, as it is far
faster
to view specimens when held in the hands of the observer. To hold specimens, pick up the head of the
pin using the thumb and forefinger of your dominant hand. This allows you to easily spin the specimen around
the axis of the pin. The point of the pin i
s then either lightly pressed against the middle or forefinger of the
other hand, or held between the thumb and forefinger.

It is important to place the bottom sides of your hands on the base of the microscope; this stabilizes the hand
so the specimen is held steadily even under high magnification. With hands in place, the specimen can be
quickly and efficiently rotated in all

directions while the observer looks into the microscope. To take full
advantage of this, the focal plane of the microscope should be raised such that the specimen is roughly in
focus (usually about 3 inches above the base of the microscope), when the han
ds are in place. Once this focus
is set on the microscope, it is never moved again, as any change in focus is accomplished by moving the
specimen rather than moving the focus knob. If the magnification level needs to be changed, the hand holding
the head

of the pin can retain the specimen while the other hand changes the magnification without having
the eyes leave the oculars.

The final part of microscope setup is to adjust your chair or the table holding the microscope such that you do
not have to bend

or strain your body to look into the microscope.

Acknowledgements: John Ascher, Harold Ikerd, Gretchen LeBuhn, Jack Neff, and Karen Wetherill had valuable
additions to this section.

Gary Alpert put us on to a clever specimen holder made fro
m a ping po
ng ball filled with P
laster of
P
aris.



You place the plaster filled ball in a large heavy washer of some kind and like a track ball you can swivel it
around to get the best look at your bee.



While not a fan of using platforms to ID bees, this is useful
for some circumstances and beats all other stands
hands down.



General steps



1. Buy a ping pong ball


2. Drill small hole in said ball


3. Mix a fairly liquid batch of
P
laster of Paris


4. Quickly transfer plaster of P
aris to ping pong ball using a syri
nge or eye dropper


5. Wash equipment immediately


6. Wait for pp ball to dry


7. Drill a small hold in plaster


8. Plug with clay


9. Optional: paint photographer gray


10. Set ball in washer


11. Put specimen in clay


12. Pivot as desired

The Bee Bowl

Tr
ap


Bee bowls are small colored plastic bowls or cups that are filled with soapy water. Bees are attracted to these
colors, fly into the water, and drown. Originally meat trays (a.k.a. pan trap) and 12 oz. salad bowls were used.
Field experience and expe
riments have demonstrated that bowl size is not necessarily correlated with capture
rate (see
http://online.sfsu.edu/~beeplot/

for several reports that document those results, or contact Sam
Droege and Gretc
hen LeBuhn for unpublished experiments on such).

Several manufacturers make such cups, but Solo is the line that most people have experience with. These cups
are usually
translucent,

which is not at all attractive to bees. However, the Solo 3.25 oz. cup is steep sided,
stable on the ground and does come in white (model number: p325w
-
0007). This particular model works well
because plain white is highly attractive to bees and it also
provides a nice base color when painting flourescent
blue or yellow.

For travel, the Solo 0.75 oz. and 2 oz. cups are nice sizes to carry in your luggage as they minimize water use.
However, they will lose water very quickly in hot, low humidity environ
ments. The 1 oz. cups are steeper
-
sided and narrow (and therefore more unstable), however, this model may be worth investigating for use in
desert areas. Surprisingly, loss or upsetting by the wind is rarely an issue with bee bowls.

The white cups usuall
y
need to be ordered by the case from a local Solo distributor (that means about 2500
cups).

Translucent models
are widely available and very inexpensive online. Solo distributors can be located
by calling 1
-
800
-
FOR
-
CUPS. The solo product line catalog i
s online and can be viewed at:
www.solocups.com
.
The price for a case of the white bowls is usually in the range of $50 to $85. Do a Google search on the

model
number and see what you can find.
Denny Johnson locat
ed a source at
www.cometsupply.com

that
,

as of the
writing of this version of the manual
,

are still available
.
See the bottom of the next section for a source of pre
-
painted bowls.

Painting Bowls



Prior to usin
g
soufflé cups, colored plastic bowls from party stores or other sources were
used to capture bees. The usual colors were yellow, white, light blue, and dark blue. Those worked well, but
fluorescent yellow and fluorescent blue were found to be much more
effective in the East (and field
experience indicates the same to be true in the West). However, note that Laurence Packer has found that
cactus bees, especially
Macrotera
, seem to be attracted to dark blue and even red colored bowls (red bowls
attracted absolutely zero bees in the East). He didn't compare these with fluorescent colors, but both those
colors collected more M. texana than did either white or yellow. A lite
rature is accumulating that indicates
that there are individual species preferences in bowl color and that these preferences appear to shift
regionally and perhaps even seasonally.

Commercial fluorescent spray and brush paints vary in their color characte
ristics and availability by brand and
location
.

In 2004, we experimented with some different formulations and found a fluorescent combination
from the East Coast Guerra Paint and Pigment that works better than the system we had earlier tried. The
liquid
pigments mix much more readily than the dry pigments and their base paint sticks well to plastic. When
ordering from Guerra (212
-
529
-
0628) specify:



Silica Flat



Yellow Fluorescent



Blue Fluorescent

Jody has been the person we have worked with.

You can ord
er online at:

http://www.guerrapaint.com/tandc.html

To get to the fluorescent pigments, click on “Search By Group”. Run the scroll bar down to the bottom and
click on “Flourescent.” Choose “Flourescent

Blue” or Flourescent Yellow” in the size and quantity you desire.

To get to the silica flat, click on “Search By Type” and choose “Binder” from the list. Choose the size and

amount of “Silica Flat” you need.

The ratio is 16 oz. of dye to 1 gallon of Sili
ca Flat Paint. You can mix it with a stick without difficulty.

For future reference, their Fluorescent (water
-
dispersed pigments) formula is:



Water







47.5%



Methocel


KMS


Thickener


Methyl Cellulose


0.45%



Defoamer


Drew
-
647





0.80%



Tamol

731


Dispersant (soap)




1.25%



Flourescent Pigment





50.0%

The formula for the Silica Flat Acrylic Latex Paint is:



Acrylic
-
Latex



Calcium Carbonate



Kaolin


Clay



Tanium Dioxide (I think this should be Titanium Dioxide)

No percentages were given and t
hese are only listed as the major components; there are likely to be
surfactants and other things in here as well. The carrier of the dye is not as important as the dye itself.

You can purchase prepainted fluorescent blue or yellow 3.25 ounce soufflé cups

from New Horizons Support
Services (
http://www.nhssi.org/
) which uses developmentally disabled workers to paint the bowls. They also
sell unpainted white bowls. Email your queries to
Cynt
hia Swift
-
King (
cking@nhssi.org
) or call the number at
the web site listed above.